We believe this site might serve you best:

United States

Select a Different Country and Language

Promega's Cookie Policy

Our website uses functional cookies that do not collect any personal information or track your browsing activity. When you select your country, you agree that we can place these functional cookies on your device.

Epigenetics refers to heritable changes in gene expression that arise from changes in
chromosomes without alteration of DNA sequence. These changes occur throughout all
stages of development or in response to environmental factors such as exposure to toxins
or chronic stress and are implicated in diseases such as cancer. Epigenetic mechanisms
of gene regulation, which collectively make up the epigenome, include modifications to
DNA and histone components of nucleosomes as well as expression of noncoding RNAs
(ncRNAs). These modifications can affect gene accessibility to DNA-binding and
regulatory proteins such as methyl-CpG-binding proteins, transcription factors, RNA
polymerase II and other components of the transcriptional machinery, ultimately altering
transcription patterns, often in tissue- and cell-specific ways. A schematic diagram
showing the most well characterized epigenetic modifications are shown in Figure 16.1.

In vertebrates, DNA methylation occurs on the 5C position of cytosine residues to
yield 5-methylcytidine. This occurs almost exclusively within CpG dinucleotides,
although nonCpG methylation does occur in plants (primarily CpNpG and CpHpH methylation,
where H = A,T,C) and to a lesser extent, mammals. Other forms of cytosine exist,
including 5-hydroxymethylcytosine, 5-formylcytosine and 5-carboxylcytosine (Kriaucionis
and Heintz, 2009; Ito et al. 2011; Pfaffeneder et
al. 2011), which may be intermediates in a pathway for DNA demethylation.

In mammalian genomes, approximately 70–80% of CpG dinucleotides are methylated.
However, stretches of CpG-rich sequences with low levels of DNA methylation, known as
CpG islands, exist (reviewed in Blackledge and Klose, 2011; Deaton and Bird, 2011). DNA
methylation is typically associated with epigenetic gene repression, and many targets of
de novo DNA methylation during differentiation are promoters of stem cell- and
germline-specific genes (Weber et al. 2007; Mohn et
al. 2008; Farthing et al. 2008). DNA methylation also
recruits methyl-CpG-binding proteins, which recruit additional proteins that add
silencing modifications to neighboring histones. This coordination between DNA
methylation and silencing histone marks leads to compaction of chromatin and gene
repression.

CpG islands (CGIs) make up only 0.7% of the human genome but contain 7% of the CpG
dinucleotides. CpG islands often are highly enriched at gene promoters, and
approximately 60% of all mammalian gene promoters are CpG-rich. CpG islands are
typically unmethylated, open regions of DNA with low nucleosome occupancy. As such,
CpG islands promote relaxed chromatin structure that favors active transcription,
known as euchromatin, and increases accessibility of RNA polymerase II and other
components of the basal transcription machinery to the transcription start site. Most
CGI promoters have heterogeneous transcription start sites and lack TATA boxes, so
transcription factors with CpG in their recognition sites, such as SP1, can help
recruit TATA-binding protein to promoters without TATA boxes. Without additional
regulatory signals, transcription from CGI promoters results in nonproductive,
bidirectional cycles of initiation and premature termination. The regulatory signals
required for the transition from this nonproductive state to productive, directional
synthesis of full-length transcripts are not yet well characterized.

The mechanisms that keep CpG islands free of methylation appear to involve binding
of transcription factors and other transcriptional machinery or the act of
transcription itself. However, CpG islands can become hypermethylated (Meissner
et al. 2008; Mohn et al. 2008) to
silence specific genes during cellular differentiation, genomic imprinting and X
chromosome inactivation.

The methylation status of a DNA sequence can be determined using a variety of
techniques such as the use of restriction enzymes (REs), which recognize short DNA
sequences and cleave double-stranded DNA at specific sites within or adjacent to
these sequences. Some REs are sensitive to methylation and will not cleave DNA if a
cytosine in their recognition sites is methylated, while other REs are insensitive to
methylation. The methylation-sensitive RE HpaII was used in early epigenetics studies
to determine that 55–70% of all HpaII sites (5′-CCGG-3′) are
methylated in the mammalian genome (Bird, 1980; Bestor et al.
1984) and to identify CpG-rich, hypomethylated DNA regions [known as HpaII tiny
fragments (HTFs); Bird, 1986].

A list of REs that are sensitive to CpG and CpNpGp methylation can be found in the
Technical Reference section of the Promega web site

Pairs of isoschizomers where one RE is insensitive to methylation and the other is
sensitive (Table 16.1) are often used to query methylation status. DNA fragments
generated by a methylation-sensitive isoschizomer will differ in size from fragments
generated by a methylation-insensitive isoschizomer. The extent of cytosine
methylation can be estimated by calculating the ratio of the different DNA fragments.

Bisulfite sequencing refers to techniques that assess DNA methylation through
bisulfite conversion, which converts unmethylated cytosine residues to uracil
residues. Methylated cytosine residues remain unmodified (Frommer et
al. 1992). The target DNA is purified, alkaline- or heat-denatured,
treated with sodium bisulfite, cleaned up, treated with alkaline, then cleaned up
again to remove salts and other components that can inhibit downstream applications.
DNA purification kits, such as the Wizard® SV Gel
and PCR Clean-Up System (Cat.# A9281), are commonly
used for this purpose. After bisulfite conversion and DNA cleanup, the DNA is
amplified by whole genome PCR, and the amplified products are analyzed using a
technique that distinguishes products derived from unmethyled DNA, which contain
thymine residues, from products derived from methylated DNA, which contain cytosine
residues. These techniques include pyrosequencing, methylation-specific PCR,
methylation-sensitive single-strand conformation analysis (MS-SSCA; Bianco
et al. 1999), high-resolution melting analysis (Wojdacz and
Dobrovic, 2007), methyl cytosine immunoprecipitation (mCIP; Zhang et
al. 2006), bisulfite methylation profiling (BiMP; Reinders et
al. 2008) and MALDI-TOF mass spectrometry (Schatz et
al. 2006). For high-throughput analysis, bisulfite-treated DNA can be
analyzed using microarrays with two sets of oligonucleotide probes, one of which is
complementary to cytosine-containing DNA and the other complementary to
thymine-containing DNA.

Typical bisulfite conversion protocols involve long incubation times under harsh
conditions, resulting in highly fragmented DNA. Promega offers the
MethylEdge™ Bisulfite Conversion System (Cat.#
N1301), which results in efficient DNA conversion and recovery with
reduced template fragmentation using a protocol that can be completed in less than
two hours, including desulphonation and cleanup. The MethylEdge™ Bisulfite
Conversion System does not require an additional cleanup kit.

Additional Resources for Bisulfite Conversion

Technical Bulletins and Manuals

The firefly luciferase reporter protein (Fluc) can be used to assess DNA
methylation at the genome level or at specific DNA sequences. Researchers have
developed split-luciferase biosensors composed of two fusion proteins: a DNA-binding
domain fused to the N-terminal portion of Fluc, and a second DNA-binding domain fused
to the Fluc C-terminus (Badran et al. 2011). To assess levels of
global DNA methylation, both fusion proteins are constructed using the DNA-binding
domain of a methyl-CpG-binding domain protein such as MBD2, which has a 100-fold
preference for methylated CpG sites over unmethylated CpG sites. The fusion proteins
are expressed in a cell-free expression system, then incubated with the target DNA to
allow DNA binding. If multiple methylated CpG sites exist in proximity, the
N-terminal and C-terminal portions of Fluc will interact (Figure 16.2). The level of
restored Fluc activity is measured using a firefly luciferase assay, such as the
Steady-Glo® or
Dual-Glo® Luciferase Assay System, and
luminescence levels are indicative of DNA methylation levels throughout the genome.
To measure site-specific DNA methylation levels, the N-terminus of Fluc is coupled to
the MBD DNA-binding domain, but the C-terminus is coupled to a sequence-specific
DNA-binding domain (Porter et al. 2008).

Figure 16.2. A schematic diagram showing the use of a split-luciferase biosensor to
assess DNA methylation at the genome level.To assess DNA methylation at a specific DNA sequence, one of the
methyl-CpG-binding domains (MBD) is replaced with a sequence-specific
DNA-binding domain. N-luc is the N-terminal portion of firefly luciferase;
C-luc is the C-terminal portion of firefly luciferase.

Epigenetic gene regulation also is controlled by changes in histones that make up the
nucleosome and histone modification. Canonical nucleosomes are octamers that consist of
H2A, H2B, H3 and H4 proteins. However, there are several histone variants that can vary
by a small number of amino acids or include large insertions (reviewed in Sarma and
Reinberg, 2005). Often these histone variants are found at specific locations within the
chromatin or are used to demarcate boundaries between heterochromatin and euchromatin
regions.

The majority of histone-mediated regulation stems from histone modification, most
often modification of the exposed amino termini of histones protruding from the
nucleosome core. The predominant histone modifications include acetylation, methylation,
phosphorylation, ubiquitination and sumoylation, with thousands of potential
combinations of modifications within a single nucleosome. Of these, histone acetylation
and methylation are the best understood, and some general trends have been observed.
Trimethylation of histone H3, specifically the lysine at position 4 (H3K4me3), is a mark
associated with transcriptionally active chromatin, whereas H3K27me3 leads to compact
chromatin, which represses gene expression. The term “histone code” is used to describe
how different combinations of histone modifications affect transcription levels.

Identification of proteins that read, write or erase these marks is critical to help
unravel the complexities of epigenetic regulation. Chromatin immunoprecipitation (ChIP)
is a powerful assay to identify proteins that bind to chromatin and map protein binding
throughout the genome using techniques such as microarray analysis or high-throughput
sequencing.

In ChIP analysis, protein:protein and protein:DNA complexes are crosslinked,
immunoprecipitated using an antibody against the protein of interest and purified. The
DNA sequence of interest then is amplified from the immunoprecipitated material using
PCR. Alternatively, the immunoprecipitated DNA can be sequenced (ChIP-seq) or analyzed
using microarrays (ChIP-chip) to identify target sequences.

Figure 16.3. Schematic diagram of the HaloCHIP™ System.

One challenge of the traditional ChIP method is the availability of specific
antibodies that recognize crosslinked epitopes. To overcome the need for suitable
antibodies, Promega scientists developed the HaloCHIP™ System (Figure 16.3).
This system takes advantage of the HaloTag® protein,
which is a mutated hydrolase (Los et al. 2005; Los and Wood, 2007;
Los et al. 2008; Hartzell et al. 2009) that
catalyzes a covalent attachment to a variety of ligands, including a resin-based ligand
for immobilization. This tag can be fused to any protein; for ChIP, the DNA-binding
protein of interest is fused to the HaloTag® protein
by cloning the protein-coding region into a HaloTag®
vector. The recombinant construct is transfected into cells for stable or transient
expression, then cells are treated with formaldehyde to induce covalent protein:DNA and
protein:protein crosslinks, lysed and sonicated to shear the DNA into smaller fragments.
The crosslinked complexes are captured directly from the lysate through covalent binding
of the HaloTag® moiety to the HaloLink™
Resin. Covalent binding allows more extensive and stringent washing than is possible
with noncovalent interactions, resulting in reduced background and increased
signal-to-noise ratio. Subsequent heating of the purified complexes reverses the
crosslinks and releases captured DNA fragments, which can be purified and analyzed using
PCR, sequencing or microarray analysis. For more information, see the
HaloCHIP™ System Technical Manual
#TM075.

Online Tools

Acetylation of a lysine residue neutralizes a positive charge on a histone
protein, reducing the electrostatic interaction with negatively charged DNA. This
reduction in affinity leads to increased accessibility of the DNA to protein
complexes, which can lead to increased gene expression. In addition, lysine
acetylation can recruit nucleosome-remodeling complexes, such as Swi2/Snf2, via their
bromodomains to promote and maintain euchromatin structure (reviewed in Bernstein
et al. 2007). However, the factors controlling gene
expression are complex, and histone acetylation also can lead to reduced gene
expression through indirect mechanisms.

Lysine acetylation occurs on the N-terminal tails of core histones and is
controlled primarily by two enzyme families: histone acetyl transferases (HATs) and
histone deactylases (HDACs). HATs use acetyl CoA as a coenzyme to transfer an acetyl
group to the epsilon amino group of the lysine side chain. These enzymes are grouped
into three families: GNAT, p300/CBP and MYST. HDACs reverse histone acetylation and
promote gene silencing. HDACs are often components of large protein complexes and are
recruited to sites of DNA methylation by methyl DNA-binding proteins. HDACs fall into
four categories: Class I, which includes HDAC1, 2, 3 and 8; Class, II, which includes
HDAC4, 5, 6, 7, 9 and 10; Class III, which includes the NAD+-dependent sirtuins
(SIRTs); and Class IV, which includes HDAC11 (reviewed in Sun et
al. 2012).

Misregulation of HATs and HDACs often is associated with development and
progression of cancer and other diseases such as neurodegenerative disorders and
cardiovascular diseases, making these enzymes attractive therapeutic drug targets.
Many HDAC inhibitors promote cell cycle arrest at the G1/S phase, and studies have
shown that tumor cells generally are more sensitive to HDAC inhibitors than normal
cells (Johnstone, 2002). Also, HDAC inhibitors can restore the ability of animals to
recall memory that had been lost in Alzheimer’s and Parkinson’s disease models,
possibly by changing chromatin structure in neurons (Fischer et
al. 2007).

To facilitate screening of potential HDAC inhibitors, Promega offers the HDAC-Glo™ I/II Assays and Screening Systems and SIRT-Glo™ Assays and Screening Systems. The HDAC-Glo™
I/II and SIRT-Glo™ Assays are single-reagent-addition, homogeneous,
luminescent assays that measure relative activities of HDAC class I and II enzymes
and sirtuins, respectively. The HDAC-Glo™ I/II Assays use an acetylated,
live-cell-permeant, luminogenic peptide substrate that is deacetylated by HDAC
activities from cells, extracts or purified enzyme sources (Figure 16.4). The
SIRT-Glo™ Assay uses a similar substrate to detect SIRT activities from
purified enzyme sources (Figure 16.5). Deacetylation of the peptide substrate is
measured using a coupled enzymatic system in which a protease in the Developer
Reagent cleaves the deacetylated peptide from aminoluciferin, which is quantified in
a luciferase-based reaction. The HDAC-mediated luminescent signal is proportional to
enzyme activity and persistent, allowing batch processing of multiwell plates in
high-throughput screening.

Figure 16.5. The single-reagent-addition SIRT-Glo™ Assay.SIRT activity deacetylates the luminogenic SIRT-Glo™ Substrate,
making the peptide sensitive to specific proteolytic cleavage that is
mediated by the SIRT-Glo™ Developer Reagent and liberates
aminoluciferin. Free aminoluciferin then can be measured using
Ultra-Glo™ firefly luciferase to produce stable, persistent light
emission. Z represents an amino-terminal blocking group that protects the
substrate from nonspecific cleavage. XXXLysine is a SIRT-optimized amino
acid sequence based on a consensus sequence derived from p53 (Abraham
et al. 2000).

Prepare initial dilutions of the putative HDAC inhibitor and known HDAC
inhibitor Trichostatin A as described in the
HDAC-Glo™ I/II Assay and Screening
System Technical Manual #TM335. Add only HDAC-Glo™ I/II Buffer to the no-inhibitor
and no-HDAC control wells.

Dilute the HDAC enzyme source using HDAC-Glo™ I/II Buffer to the
desired concentration. If using the HeLa Nuclear Extract supplied with the
HDAC-Glo™ Screening System, dilute the extract 1:3,000.

Dispense the HDAC enzyme source to each well of inhibitor dilutions prepared
in Step 1 and no-inhibitor controls. Add HDAC-Glo™ I/II Buffer to the
no-HDAC controls. (Dispense 50µl for 96-well plates, 10μl for 384-well plates
or 2.5μl for 1536-well plates.)

Mix the plate at room temperature for 30–60 seconds using an orbital shaker
at 500–700rpm to ensure homogeneity.

Incubate enzyme/inhibitor mixes at room temperature for at least 30 minutes
but not longer than approximately 2 hours.

Prepare the HDAC-Glo™ I/II Reagent as described in the
HDAC-Glo™ I/II Assay and Screening
System Technical Manual #TM335.

Add an equal volume of HDAC-Glo™ I/II Reagent to each assay well
(100μl for 96-well, 20μl for 384-well or 5μl for 1536-well plates).

Mix the plate at room temperature for 30–60 seconds using an orbital shaker
at 500–700rpm to ensure homogeneity.

Additional Resources for the HDAC-Glo™ I/II and SIRT-Glo™
Assays and Screening Systems

Technical Bulletins and Manuals

Histone methylation occurs at lysine residues, which can be mono-, di- or
trimethylated, and arginine residues, which can be mono- or dimethylated. Histone
methylation is catalyzed by protein lysine methyltransferases (PKMTs) and protein
arginine methyltransferases (PRMTs) but can be reversed by protein demethylases. To
date, researchers have identified >30 demethylating enzymes, >50
protein lysine methyltransferases and >10 protein arginine methyltransferases,
suggesting that protein methylation is a dynamic and complex process (Janzen
et al. 2010). Histone methylation has different effects on
transcriptional activity, depending on the number of methyl groups and position of
the amino acid being modified. In general, the H3K9me1 mark is activating, whereas
H3K9me2 and H3K9me3 are repressive; H3K4me3 and H3K36me3 are associated with active
chromatin, whereas H3K9me3, H3K27me3, H3K36me2 and H4K20me1 often are found in
transcriptionally repressed heterochromatin.

The downstream effects of histone methylation are largely determined by proteins
that bind to the modified histones. For example, H3K9me3 acts as a binding site for
heterochromatin protein 1 (HP1), which then can recruit histone methyltransferases,
histone deacetylases and other proteins that affect chromatin structure. H3K4me3
recruits proteins that promote euchromatin, whereas H3K9me1, H3K9me2 and H3K27me3
interact with proteins that promote heterochromatin. Two such groups of proteins are
the polycomb group (PcG) proteins and their antagonists, the trithorax (trxG) group
proteins, which were first identified as regulators of hox gene
expression in Drosophila (Schwartz and Pirrotta, 2008). More
recent studies have shown that related proteins exist in mammals and plants. PcG
proteins repress transcription; trxG proteins activate transcription. Some PcG and
trxG proteins possess histone methyltransferase activity and can modify histones
directly, while others bind to and interpret histone modifications.

In embryonic stem (ES) cells, CpG islands that are regulated by PcG proteins often
are “bivalent” in that they retain the permissive H3K36me2-depleted and
H3K4me3-enriched environment but also exhibit H3K27me3. Genes with bivalent promoters
often are actively silenced in ES cells but lose the repressive H3K27me3 mark while
retaining the activating H3K4me3 mark later during differentiation.

Histones can be phosphorylated on serine, threonine and tyrosine residues. Many of
the serine and threonine phosphorylation events play a role in DNA repair or DNA
condensation, segregation and decondensation during mitosis, but some are involved in
epigenetic regulation of transcription, including H3T3ph, H3T6ph, H3T11ph, H2.AS1ph,
H3S10ph and H4S41ph (reviewed in Pérez-Cadahía et al. 2010).
H3S10ph is one of the best characterized of these histone modifications. In addition
to its DNA-restructuring responsibilities during mitosis, H3S10ph seems important for
chromatin decondensation associated with transcriptional activation of target genes.
H3S10ph recruits chromatin-modifying enzymes and chromatin-remodeling complexes and
prevents binding of HP1 to neighboring H3K9me3 marks at the onset of mitosis.
H3S10ph, as well as H3T3ph and H3T11ph, can block binding of DNMT3a to H3, reducing
methylation of nearby chromatin (Zhang et al. 2010).

Histones contain many highly conserved tyrosine residues, many of which can be
phosphorylated. Phosphorylation of H3Y99 is critical for polyubiquitination and
subsequent proteolysis of excess histones, which can increase a cell’s sensitivity to
DNA-damaging agents, cause genomic instability and induce apoptosis. Another tyrosine
residue, H3Y41, is important in chromatin structure and oncogenesis. In human
hematopoietic cell lines, phosphorylation of H3Y41 by Janus kinase 2 (JAK2)
destabilizes binding of HP1α to histone H3 (Dawson et
al. 2009), leading to a more open chromatin structure around certain gene
promoters such as leukemia oncogene LMO2, which can trigger
oncogenesis in hematopoietic cells. Overexpression or aberrant activation of JAK2
activity leads to higher levels of H3Y41, loss of HP1α binding and higher
expression of LMO2.

Promega offers a number of kinase enzyme systems to monitor the activity or
identify inhibitors of different kinases involved in histone phosphorylation,
including IKKα, PIM1, RSK2 and several cyclin-dependent kinases (CDKs) such
as CDK1, CDK2 and CDK5. These luminescent assays convert ADP produced by these
kinases to ATP, which is then converted to light by Ultra-Glo™ Luciferase.
The resulting luminescent signal positively correlates with ADP amount and kinase
activity. An example protocol is provided below. For a list of available Kinase
Enzyme Systems, refer to the Human Kinome
chart.

kinase enzyme prepared in 1X Kinase Buffer A at 2.5 times the desired
final concentration (We recommend an amount that will convert 5–10% of the
ATP to ADP.)

CDK1 inhibitor of interest

5% DMSO prepared in 1X Kinase Buffer A, as a no-inhibitor control

water

luminometer capable of reading multiwell plates

plate shaker

Reagent Preparation

Thaw Kinase Detection Buffer at room temperature. If a precipitate is
present, incubate the buffer at 37°C with constant swirling for 15 minutes.
Alternatively, remove the precipitate from the Kinase Detection Buffer by
carefully pipetting the supernatant from the bottle.

Conjugation of ubiquitin, a 76-amino acid protein, to lysine residues of histone
proteins can affect transcription activity as well as nucleosome stability and, as a
result, gene accessibility. The consequences of histone ubiquitination depend on the
histone substrate and degree of ubiquitination (reviewed by Weake and Workman, 2008).
Mono-ubiquitination of histone H2A (H2Aub1) is often considered a repressive mark,
while H2B mono-ubiquitination can play a role in both transcriptional activation and
silencing. In addition, there is evidence of cross-talk between histone
ubiquitination and other forms of histone modification. For example, ubiquitinated
H2B has been identified as a docking site for the COMPASS protein complex
(Chandrasekharan et al. 2010), which includes the histone
methyltransferase responsible for H3K4 methylation. Also, H2Aub, but not H2A,
specifically represses di- and trimethylation of H3K4, and ubiquitin-specific
protease 21 (USP21) relieves this repression (Nakagawa et al.
2008).

To measure the activity of deconjugating enzymes such as deubiquitinating (DUB),
deSUMOylating (SENP) and deneddylating (NEDP) proteases, Promega offers the
homogeneous, bioluminescent DUB-Glo™ Protease Assay (SUB/SENP/NEDP)
(Cat.# G6260). This assay uses a luminogenic
substrate that contains the C-terminal pentapeptide of ubiquitin:
Z-RLRGG-aminoluciferin. Upon addition of the DUB-Glo™ Reagent to the test
sample, the substrate is cleaved to form the luciferase substrate aminoluciferin,
resulting in a glow-type luminescent signal that is proportional to the amount of
DUB, SENP and NEDP1 activity present.

Additional Resources for Histone Ubiquitination

Technical Bulletins and Manuals

Another post-translational modification that plays an important role in epigenetic
regulation is sumoylation, the addition of the small ubiquitin-related modifier SUMO
(reviewed in Ouyang and Gill, 2009). This modification can stabilize proteins, alter
subcellular localization, affect enzyme activity and mediate interactions with other
proteins. Many transcription factors and cofactors can be sumoylated, which is
generally indicative of transcription repression. In Drosophila,
the sumoylated form of Sp3 recruits the polycomb protein Sfmbt (Steilow et
al. 2008a) and HP1α, β and γ (Steilow
et al. 2008b; Seeler et al. 2001) to
repress transcription.

Many histone-modifying enzymes, nucleosome-remodeling complexes and their
associated enzyme cofactors contain one or more SUMO interaction motifs (SIMs). This
motif allows these proteins to interact with sumoylated transcription factors and
cofactors, which can direct these enzymes to specific promoters. Two such groups of
histone-modifying enzymes recruited by SUMO are histone deacetylases, which decrease
histone acetylation at the target promoter, and histone demethylases such as
lysine-specific demethylase 1, which catalyzes the removal of methyl groups from
H3K4.

Many important regulatory proteins contain domains that bind to modified residues,
including plant homeobox domain (PHD) fingers, bromodomains, chromatin organization
modifier (chromo) domains, WD40 repeat and tudor domains. Transcription-friendly H3K4me3
acts as a binding site for effector proteins that contain a PHD finger, such as
nucleosome remodeling factor (NURF) and the ING4-containing histone acetyltransferase
complex. The H3K36me2 modification, which interferes with transcription initiation, acts
as a binding site for the chromodomain of the RPD3S histone deacetylase complex.

Histone modifications also can act in cooperation. The specific combination of
histone modifications at a particular site often determines which protein complexes and
accessory proteins are recruited to activate or repress transcription directly, catalyze
additional histone modifications or recruit other histone-modifying proteins. This
cooperativity can be explained, at least in part, by the fact that these proteins can
contain one or more modified-histone-binding domains. For example, the TFIID protein
complex contains both a PHD finger and bromodomain and so binds more strongly to H3K4me3
marks near acetylated H3K9 and H3K14 residues.

The absence of one of these domains through gene mutation or rearrangement can cause
serious lapses in gene regulation and diseases such as cancer. Recently, researchers
characterized a translocation involving histone demethylase KDM5A that resulted in
fusion of the H3K4me3-binding PHD finger of KDM5A to the transcriptional activator
NUP98, a common leukemia translocation partner, in an acute myeloid leukemia patient
(Islam et al. 2011). Similarly, mixed lineage leukemia (MLL) family
members, which act as histone methyltransferases, are involved in translocations in
MLL.

There is increasing evidence that expression of noncoding RNAs, such as microRNAs,
small RNAs and large RNAs, play a role in epigenetic gene regulation (reviewed in Costa,
2008; Chuang and Jones, 2007). Noncoding RNAs can direct both cytosine methylation and
histone modification to silence DNA repeats in the genome. For example, a class of
29-nucleotide RNAs that was first discovered through their interaction with the
spermatogenesis-specific PIWI protein (piwi-interacting RNAs; piRNAs) map to repetitive
DNA sequences and are important for silencing short interspersed elements (SINEs), long
interspersed elements (LINEs) and long terminal repeat (LTR) retrotransposons.

Several noncoding RNAs also are implicated in X chromosome inactivation. The 17kb
X-inactive specific transcript (XIST) RNA binds and coats the inactive X chromosome and
forms complexes that modify chromatin structure to suppress transcription. XIST levels
are controlled by another large noncoding RNA, TSIX, which is transcribed from the
strand opposite of XIST. Noncoding RNAs are involved in genomic imprinting via a similar
mechanism (reviewed by Koerner et al. 2009).

Maintenance and inheritance of epigenetic marks during cell division is critical to
maintain a committed cell lineage and cellular phenotype in progeny cells, and set a
memory of transcriptional status. The transmission of epigenetic information through
multiple cell divisions involves many of the mechanisms discussed in this chapter: DNA
methylation, histone modification, histone variants and expression of noncoding RNAs
(reviewed in Zaidi et al. 2011). These same mechanisms govern the
inheritance of epimutations, which can lead to changes in chromatin structure and
transcription levels of genes important to diseases such as cancer and imprinting
disorders.

Aberrant regulation of epigenetic mechanisms can result in genomic imprinting
disorders, such as Angelman syndrome and Prader-Willi syndrome, and may contribute to
the heritability of many forms of cancer, asthma, Alzheimer’s disease and autoimmune
diseases such as systemic lupus erythematosus, rheumatoid arthritis and multiple
sclerosis (reviewed in Hirst and Marra, 2009; Hewagama and Richardson, 2009; Handel
et al. 2010). Epimutations can interfere with epigenetic
regulation at many levels, including DNA methylation, histone modification and noncoding
RNAs. Some epimutations are inherited, but many accumulate due to environmental factors
or with age. For example, even though monozygotic twins are epigenetically
indistinguishable at birth, their patterns of DNA methylation and histone acetylation
can differ dramatically as they age (Fraga et al. 2005).

Scientists at Your Service

We offer a range of services to help you succeed using Promega technologies. From product training to set up of automated systems and development of custom applications—our scientific support goes beyond the basics.